Abstract

Investigating how the protein machinery in cells functions is fundamental to biological enquiry. Technology that can identify
and quantitate a multitude of unknown proteins can facilitate such investigations. Although techniques such as Western blotting
are able to provide limited but useful information about a known protein if a suitable antibody is available, mass spectrometry
provides a platform for the large‐scale identification and quantitation of the many unknown proteins in a typical sample.
In particular, shotgun proteomics techniques have been used for studying global changes in whole proteomes, probing the subunit
composition of protein complexes, and for mapping post‐translational modifications. The advent of whole genome sequencing
has enabled recent advances in shotgun proteomics technology. Multidimensional protein identification technology (MudPIT),
which couples mass spectrometry and multidimensional chromatography, achieves exquisitely sensitive protein detection. There
are a variety of methods used to quantify the identified proteins, from isotopic labelling to ‘label free’ spectral counting.

Key Concepts:

‘Shotgun proteomics’ aims to identify all proteins present in a sample using mass spectrometry to make measurements that depend
on a protein's mass.

Before mass spectrometry, proteins in a sample are digested with a protease to create a complex peptide mixture for analysis.

Peptides between approximately 10 and 30 amino acids long can generate characteristic fragmentation patterns that can be used
to identify their sequences.

Chromatography is used to enrich peptides of the same kind; if enough of these enter the mass spectrometer concurrently, they
can be characterised.

These homogeneous populations of peptides are isolated and fragmented inside the mass spectrometer; as these fragments are
detected, MS/MS spectra are generated.

The patterns described by these MS/MS spectra are peptide ‘fingerprints’ which can be matched to predicted spectral patterns;
genomic nucleic acid sequences are used to predict these spectral patterns.

Protein quantitation is possible; strategies used include counting the number of MS/MS spectra per protein obtained during
the analysis (label free), or labelling proteins/peptides from two samples with different isotopes.

MudPIT workflow. (a) Proteins extracted from cells are digested into peptides, which are then fractionated using a combination
of ion exchange and reverse phase chromatography. The peptides are sprayed directly into the mass spectrometer. After the
MudPIT ‘run,’ experimentally generated MS/MS scans are analysed computationally and peptides/proteins identified. (b) Shotgun
proteomics methods experimentally characterise short peptides rather than intact proteins. Such peptides dissociate inside
the mass analyser in a predictable way to generate N‐terminal (y) or C‐terminal (b) ions. The mass to charge ratio of these ions results in a pattern in the MS/MS spectra that depends on the peptide
sequence.

Figure 2.

Microcapillary chromatography. (a) Peptides are separated chromatographically using approximately 15 cm long silica microcapillary
columns. The tip of the column is pulled to approximately 5 μm and columns packed with approximately 5 μm diameter chromatography
resin. (b) Columns are packed with three regions of reverse phase (RP) or strong cation exchange (SCX) resin. The column is connected to an HPLC pump and the peptides are eluted across a approximately 5 mm gap into the mass spectrometer. A potential difference of approximately
2 kV between the buffer in the column and the mass spectrometer results in solvent free peptide ions (electrospray ionisation
(ESI)).

Figure 3.

Resolving peptides with multidimensional chromatography. (a) During the initial 120 min chromatography step, a gradient of
the organic solvent acetonitrile is used to disrupt hydrophobic interactions between peptides and the first region of reverse
phase (RP) resin. Peptides can then bind to the strong cation exchange (SCX) resin. (b) In subsequent 120 min steps a salt ‘bump’ is used to transfer some peptides to the second region of reverse phase
resin. These peptides are then gradually eluted into the mass spectrometer with an acetonitrile gradient. The salt concentration
is increased in each successive salt bump.

Figure 4.

Generating MS/MS spectra. (a) An initial ‘MS’ scan is used to survey the different species of peptides entering the mass spectrometer.
A sample of peptides is trapped and then peptides are gradually ejected through slits on two of the poles of the mass analyser.
Peptides with lower m/q (mass/charge) values are ejected first. Data from this MS scan is used to select a particular m/q
value for the subsequent ‘MS/MS’ analysis. (b) A fraction of a second after the MS scan, more peptides are trapped. Peptides
with a chosen value of m/q (the red peptides) are then isolated. These (red) peptides collide with helium atoms in the mass
analyser, absorb energy and dissociate. The fragments with different m/q are gradually ejected for detection as before. The
resulting MS/MS spectrum can be used to identify the peptide. The data from each MS scan can be used to collect several MS/MS
spectra.

Figure 5.

Labelling samples for quantitation. Proteins can be labelled with either heavy or light amino acids in the growth medium during
culture (SILAC). After lysis, proteins are mixed prior to analysis. Peaks from peptides with either heavy or light amino acids are slightly
offset in the MS spectrum; the relative intensity of the MS peaks can be used for quantitating the relative abundances of
the parent proteins in the two samples. Alternative techniques include labelling proteins (ICAT) or peptides from digested proteins (iTRAQ).